1. Field of the Invention
The present invention relates generally to analytical methods where fluorescence spectroscopy is used to analyze small numbers of molecules that are present in a relatively small detection volume or zone. More particularly, the present invention involves determining a wide variety of physical and chemical properties of one or more molecules by rapidly modulating the wavelength, intensity and/or polarization of beams of laser energy to excite fluorophores that are attached either to the molecule of interest or a molecule that interacts with the molecule of interest. The emission profile of the fluorophore is used to determine useful information about various properties of the labeled and/or non-labeled molecules including molecular interactions between the molecules
2. Description of Related Art
The publications and other reference materials referred to herein to describe the background of the invention and to provide additional detail regarding its practice are hereby incorporated by reference. For convenience, the reference materials are numerically referenced and grouped in the appended bibliography. The contents of these publications and other reference materials are hereby incorporated by reference.
Understanding the intricate network of interactions occurring in cells allows probing of the mechanisms that control cell growth, maintenance and disease/death. Such interactions include protein-protein, protein-DNA, nucleic acid-nucleic acid, antibody-antigen, receptor-ligand, protein-drug and aggregation-inducing protein-protein interactions. Rapid, reliable and inexpensive methods that can characterize the multitude of the existing interactions is a core technology for proteomics, the scientific domain associated with the mapping of the complement of pair-wise protein-protein interactions on an organism-wide basis. Such interaction maps are necessary for the deciphering of the cell circuitry. Proteomic technologies are a prerequisite for interpreting, utilizing and leveraging the plethora of the genomic information available in the public domain. In-vitro and in-vivo screening of compound libraries against specific protein interactions will allow discovery of specific and powerful pharmaceuticals. Moreover, affordable versions of ultra-sensitive detection methods will an extremely useful tool for point-of-care diagnostics of a modem medical environment.
Several methods exist for evaluation of protein-protein interactions [1]. The yeast two-hybrid (Y2H) system, a genetic system based on the transcriptional activation of specific yeast genes [2-8] is a popular one. However, Y2H systems have limitations including complications associated with using protein fusions, incorrect protein folding, incorrect post-translational modification, potential toxicity of the proteins of interest in yeast, erratic reproducibility with large fraction of false positives and false negatives [1,3]. Additionally, since the observable of Y2H is based on DNA transcription, the Y2H method is not suited for the study of DNA transcription factors or DNA-binding proteins. Finally, use of Y2H requires several hours (due to the requirement of growing yeast cells) and cannot provide the accurate and complete characterization available for a homogeneous, solution-based, equilibrium-binding assay.
Development of optical methods that detect interactions in minute solution volumes and high-throughput screening (HTS) formats are highly desirable, since they represent rapid, inexpensive, reliable and non-invasive ways to determine binding constants, specificities, and half-lifetimes of the complexes of interacting partners.
Fluorescence detection with ultrahigh-sensitivity has led to the development of solution-based methods that study biomolecules and their interactions in small volumes (attoliter-femtoliter) and at low analyte concentrations (pM-nM). The first such method is fluorescence correlation spectroscopy (FCS), introduced to measure chemical kinetics and molecular diffusion by analysis of concentration fluctuations of a small ensemble of molecules [9]. FCS belong in a broader family of methods, often described as fluorescence-fluctuation spectroscopy (FFS). At very low concentrations, the fluctuations are manifested by well-resolved bursts of fluorescence corresponding to molecules traversing the confocal spot. These bursts are amenable to histogram-based methods. At higher concentrations, bursts are no longer resolved and the fluctuations are better analyzed by correlation-based methods.
Single-channel FCS (referring to a single emission channel) measures the autocorrelation of fluorescence fluctuations detected from an open detection volume including microfluidic flow systems and encapsulated molecules in small volumes, subjected to flow (or drift) and diffusion. The decay curve of the autocorrelation is used to identify changes in the diffusion time of a single species or changes in the relative contribution of subpopulations with different diffusion times. Single-channel FCS has been used to measure the kinetics of nucleic-acid hybridization [10,11] and of acetylcholine-receptor interaction [12]. However, since diffusion times scale proportionally with m1/3 (m is mass), doubling the mass (e.g. due to protein homodimerization) increases the diffusion time only by 20%, a change difficult to measure with single-channel FCS and to use reliably for separating subpopulations.
Oligomerization of labeled monomeric biomolecules is also detectable by the increased brightness of the oligomers (compared to the monomer). For example, the tetramer A4 is approximately 4 times brighter than the monomer A (excluding potential quenching of fluorophores). As a result, brightness-based analysis of interactions is more sensitive and robust than diffusion-time-based analysis. This property was exploited by methods that measure oligomerization using higher-order correlation and moment analysis of photon-count histograms [13]. Direct fitting of photon-count histograms with fixed time-bin width for determination of molecular brightness was introduced by Fluorescence Intensity Distribution Analysis (FIDA; [14]) and by Photon-Counting Histograms (PCH; [15]). These methods were used to analyze ligand-protein binding [16] and cleavage of hybridized DNA by restriction enzymes [14]. However, FIDA and PCH do not account for different diffusion times of different species. As a result, their extracted parameters are often skewed. Moreover, FIDA/PCH time bins are chosen to be much smaller than diffusion time (to assume that molecules are immobile during the chosen time bin), leading to complete loss of diffusion-time information for all species.
FIDA was extended to account for diffusion by plotting photon-count histograms for different time-bin widths simultaneously (Fluorescence Intensity Multiple Distribution Analysis, FIMDA [17]). Diffusion effects change the shape of the photon-count histogram, allowing global fitting of the histograms for all time-bin widths. This fitting extracts diffusion times, as well as brightness and concentration. However, FIMDA discards useful temporal information, such as the correlation between photons within a given time bin, as well as the correlation between successive time bins.
It is relatively difficult to distinguish between subpopulations in solution based on the observables available with single detection channel methods. Consider a simple equilibrium binding between two species A and B:A+B⇄ABIf one species is labeled with a fluorophore (AF), then the species AF and AFB are distinguishable only by change in diffusion time across the detection volume. Species B must be much larger than species A for the difference to be detectable. If both A and B are labeled, then the species AFBF is easier to separate, although it remains difficult since the change results in only a 2-fold change in fluorescence. Dual-channel methods provide greater resolving power than single-channel methods. Additionally, any Forster resonance energy transfer (FRET) occurring between the fluorophores on the two species AF and BF results in quenching, which results in a complex AFBF which has two fluorophores, but not twice the brightness. With two different fluorophores detected in two detection channels, this drawback becomes an advantage, as discussed below.
To study interactions between two macromolecules using dual-channel methods, each is labeled with a fluorophore with a distinct emission wavelength rang. The fluorophores can be excited by a single- or dual-laser excitation source. If a protein A is labeled with a green-emitting fluorophore G (protein AG), and a protein B is labeled with a red-emitting fluorophore R (protein BR), association of the two proteins will yield AGBR, for which a signal in the green emission channel coincides with a signal in the red emission channel. Such detection is referred to as correlated or coincidence detection. The two distinct fluorophores can be excited using two different laser excitation wavelengths [18], two-photon excitation [19] or with a single laser in the special case of energy transfer (ET) dyes [20] or semiconductor nanocrystals [21,22]. Dual-channel methods allow more sensitive detection of molecular interactions than single-channel methods.
An interaction can be detected using cross-correlation analysis of the photon streams for the two emission channels. The autocorrelation amplitudes of each channel result from free proteins (AG and BR) and the complex AGBR, while the cross-correlation amplitude results only from the complex AGBR.
An interaction can also be monitored by changes in the relative brightness of the two channels for fixed time-bin widths (2D-FIDA, [23]). Free AG or BR emit in “green” or “red” channel respectively, while AGBR emits in both channels. This is evident after plotting the two-dimensional histogram of photon counts, where x and y axes correspond to the number of photon counts (for a fixed time-bin width) detected in “green” and “red” channel, respectively. The position of the free and bound species on the 2D-FIDA histogram species is along the axes and along the diagonal, respectively.
Photon Arrival-time Interval Distribution analysis (PAID) is an analytical method that is applicable to both single- and dual-channel formats (see WO 2004/011903A2). PAID uses fluorescence fluctuations to extract simultaneously coincidence, brightness, diffusion time, and concentration of fluorescently-labeled molecules diffusing in a confocal detection volume. PAID is based on recording photon arrival times, and plotting two-dimensional histograms of photon pairs, where one axis is the time interval between each pair of photons 1 and 2, and the second axis is the number of other photons detected in the time interval between photons 1 and 2. PAID is related to Fluorescence Correlation Spectroscopy (FCS) by a collapse of the PAID histogram onto the time interval axis. PAID extends auto-and cross-correlation FCS by measuring the brightness of fluorescent species. PAID measures brightness while retaining information on the temporal correlation of photons, and it was shown to match or exceed other FFS methods in the accuracy of separating free and bound species. PAID can be useful for detecting static and transient interactions in-vitro and in-vivo based on one-, two- or more color coincident detection of single molecules (or small ensembles) in a small confocal or wide-field detection volumes.
Single-molecule fluorescence spectroscopy (SMFS) is an ultra-sensitive optical method for detection and analysis of individual molecules. SMFS uses laser-excitation sources to probe individual diffusing or surface-immobilized fluorescent molecules and measure their fluorescence intensity, lifetime, anisotropy, and/or spectra, yielding information about molecular structure, interactions, and dynamics. During solution-based SMFS of dilute solutions of fluorescent species, single species are observed as fluorescence “bursts” that arise when the species diffuses through the detection volume. Often, the existence of an interaction is identified using Förster resonance energy transfer (FRET). If a molecule is labeled by a pair of complementary fluorescent probes (a donor, D; and an acceptor, A) in close proximity (2-10 nm), FRET can serve as a “molecular ruler”, yielding D-A distance information. FRET has been used widely for analysis of: structure and dynamics of ensembles; single-molecules (kinetics of protein and RNA folding); DNA dynamics; rotation of molecular motors; heterogeneity/dynamics of protein-DNA complexes; and single cells (using naturally-fluorescent proteins, such as GFPs and dsRed).
Dual-channel Single Molecule Fluorescence Spectroscopy (SMFS) methods have been developed for use with ratiometric observables [24-26]. The most popular ratio recovered by dual-channel SMFS is the efficiency of Förster resonance energy transfer [27-29] (also known as fluorescence resonance energy transfer), which indicates close proximity (2-10 nm) between a pair of complementary fluorescent probes, a FRET donor and a FRET acceptor [27-29]. The relationship between the FRET efficiency and distance is shown in equation 1:
                    E        =                  1                      1            +                                          (                                  R                  /                                      R                    0                                                  )                            6                                                          (        1        )            where E is the FRET efficiency, R is the donor-acceptor distance, and Ro is a constant that depends on the spectroscopic properties of the fluorophores and on the physical properties of the solution. Since the relationship of FRET and donor-acceptor distance is well-characterized, the FRET efficiency can serve as a “molecular ruler”, yielding donor-acceptor distance information. If the donor and acceptor groups are on two different molecules, existence of intermolecular FRET between donor and acceptor marks the existence of an interaction between the labeled molecules.
FRET has been used widely for analysis of structure and dynamics of ensembles [27,28]; of single-molecules (kinetics of protein and RNA folding [26,30-32], and heterogeneity/dynamics of protein-DNA complexes [33,34]); and single cells (using naturally-fluorescent proteins, such as GFPs and dsRed [35].
In the current single-molecule FRET procedures where only the donor is excited directly, there are several limitations that prevent its application to quantitative analysis of simple bimolecular interactions, such as ADonor+BAcceptor⇄ADonorBAcceptor (where ADonor is donor-labeled molecule A, and BAcceptor is acceptor-labeled molecule B). These limitations include the following:
Proximity constraint: FRET can be used only when donor-acceptor distances in the ADonorBAcceptor complex are sufficiently short (RDonor-Acceptor is less than 6-8 nm, depending on the donor-acceptor pair) to give appreciable FRET that is distinct from donor-only species. Otherwise, low-FRET ADonorBAcceptor species are indistinguishable from free Adonor species. The proximity constraint limits the ability of FRET to monitor interactions, since it is difficult to satisfy in all cases, especially for large complexes or pairs of proteins of unknown structure.
Existence of non-absorbing acceptors: “dark” states and/or photobleaching of most FRET acceptors (such as the far-red fluorophores Cy5 and Alexa647) yield species with donor-only characteristics (“donor-only” peak; [25]); the species contributing to the zero peak mask species with low FRET (corresponding to large donor-acceptor distance), leading to an apparent increase of the actual free Adonor (plus any low-FRET ADonorBAcceptor species) population.
Interference of background and impurities: Free donor-only species emit at shorter wavelength range compared to free acceptor-only species; shorter wavelength range are more prone to buffer impurities and Raman/Rayleigh scattering, decreasing the sensitivity for the identification of fluorescent species.
Inability to detect acceptor-only species: No acceptor-only species are seen either in the absence or presence of FRET, since the direct excitation of acceptor at the wavelength of the excitation of the donor is minimized to avoid crosstalk problems.
Need for corrected FRET ratio: Currently, most smFRET studies do not concentrate on obtaining accurate distances, bur rather concentrate on observing distance changes and the kinetics of such changes. A major reason for this fact is the approximate nature of the FRET-based ratio determined by the present, single-laser excitation FRET instrumentation.
Fluorescence-fluctuation spectroscopy (FFS) methods monitor interactions and dynamics by measuring timescales of fluorescence fluctuations (fluorescence correlation spectroscopy, FCS) or amplitudes of fluorescence fluctuations (such as Fluorescence Intensity Distribution Analysis, FIDA). The fluctuations result from the diffusion of limited number of fluorescent molecules through a small detection volume. Fluctuations in one or two emission ranges can be monitored with dual emission methods being more robust. Dual emission methods require two non-interacting fluorophores with distinct emission ranges. The fluorophores are excited using two different laser-excitation wavelengths or two-photon excitation sources.
Performing the dual-channel measurements with dual-laser excitation format (as with cross-correlation FCS and certain applications of 2D-FIDA) solves some of the problems with single-laser excitation. For example, acceptor-only species are now visible and species with donor and acceptor both present with no FRET are now visible. However, new problems arise that prevent its application to quantitative analysis of certain simple bimolecular interactions, such as ADonor+BAcceptor⇄ADonorBAcceptor. These problems include:
Proximity constraint: Dual laser excitation cannot distinguish well between high FRET species and acceptor only species, since both only emit in the acceptor channel.
Need for corrected FRET ratio: FRET efficiency may be determined by comparing the amount of acceptor fluorescence detected to the amount of donor fluorescence detected. This is complicated, however, by the fact that the majority of the acceptor fluorescence detected comes from direct excitation. This increases the noise on the FRET-induced signal, decreasing the accuracy of FRET efficiency extracted.